Estradiol induces heparanase-1 expression and heparan sulphate proteoglycan degradation in human endometrium.
ABSTRACT This study seeks to determine whether estrogen is able to regulate the expression of heparanase-1 (HPR1) in human endometrium.
HPR1 expression and heparan sulphate (HS) deposition in the endometrium collected in various menstrual phases were analysed by immunohistochemical and immunofluorescence staining, respectively. HPR1 expression in the endometrial cells unexposed or exposed to estradiol was analysed by using RT-PCR and luciferase reporter assay. HPR1 activity was analysed by using a novel enzyme-linked immunosorbent assay (ELISA). Cell surface HS levels were analysed by flow cytometry. Serum HPR1 activity in women receiving follicle-stimulating hormone (FSH) for IVF was measured by ELISA.
HPR1 expression was rarely detected in the endometrium in the early and mid-proliferative phases but was increased in the late proliferative phase and in the secretory phases. HPR1 expression was negatively associated with HS in the basement membrane (BM) of the endometrial glands. HPR1 gene expression, HPR1 promoter activity and HPR1 enzymatic activity were increased in the endometrial cells when exposed to 17beta-estradiol (E(2)), whereas cell surface HS levels showed a decrease which could be blocked by PI-88, an HPR1 inhibitor. Serum HPR1 levels were increased in women with moderately elevated blood estrogen levels after receiving FSH.
HPR1 is differentially expressed in the endometrium in different menstrual phases. Estrogen plays an important role in inducing HPR1 expression, subsequently leading to HS degradation on the endometrial cell surface and in the BM of the endometrium.
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ABSTRACT: Heparanase is the only known mammalian endoglycosidase capable of degrading heparan sulfate glycosaminoglycan, both in extracellular space and within the cells. It is tightly implicated in cancer progression and over the past few decades significant progress has been made in elucidating the multiple functions of heparanase in malignant tumor development, neovascularization and aggressive behavior. Notably, current data show that in addition to its well characterized role in cancer, heparanase activity may represent an important determinant in the pathogenesis of several inflammatory disorders, such as inflammatory lung injury, rheumatoid arthritis and chronic colitis. Nevertheless, the precise mode of heparanase action in inflammatory reactions remains largely unclear and recent observations suggest that heparanase can either facilitate or limit inflammatory responses, when tissue/cell-specific contextual cues may dictate an outcome of heparanase action in inflammation. In this review the involvement of heparanase in modulation of inflammatory reactions is discussed through a few illustrative examples, including neuroinflammation, sepsis-associated lung injury and inflammatory bowel disease. We also discuss possible action of the enzyme in coupling inflammation and tumorigenesis in the setting of inflammation-triggered cancer.Matrix biology: journal of the International Society for Matrix Biology 03/2013; · 3.56 Impact Factor
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ABSTRACT: Recent years have seen a growing body of evidence that enzymatic remodeling of heparan sulfate proteoglycans profoundly affects a variety of physiological and pathological processes, including inflammation, neovasvularization and tumor development. Heparanase is the sole mammalian endoglycosidase that cleaves heparan sulfate. Extensively studied in cancer progression and aggressiveness, heparanase enzyme was recently implicated in several inflammatory disorders as well. Although the precise mode of heparanase action in inflammatory reactions is still not completely understood, the fact that heparanase activity is mechanistically important both in malignancy and in inflammation argues that this enzyme is a candidate molecule linking inflammation and tumorigenesis in inflammation-associated cancers. The elucidation of the specific effects of heparanase in cancer development, particularly when inflammation is a causal factor, will accelerate the development of novel therapeutic/chemopreventive interventions and help to better define target patient populations in which heparanase-targeting therapies could be particularly beneficial. © 2013 The Authors Journal compilation © 2013 FEBS.FEBS Journal 02/2013; · 4.25 Impact Factor
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ABSTRACT: To investigate the role of heparanase-1 in laser-induced choroidal neovascularization (CNV). Experimental CNV was induced by krypton laser photocoagulation in 15 male Brown Norway rats. Fundus fluorescein angiography and histopathological examination were performed in observing the CNV development. The expression and distribution of heparanase-1 protein in the laser lesions were determined by immunohistochemistry and western blotting analysis. The success rate of laser induced CNV was approximately 75% on 3-4 weeks after laser photocoagulation. The protein levels of heparanase-1 increased significantly in the retina-choroidal complex of CNV models when compared to normal rat eyes (P<0.01). Immunostaining confirmed strong heparanase-1 expressions in all laser lesions, and it displayed to be highest at the newly formed blood vessels within the fibrovascular complex in the subretinal space. Heparanase-1 is closely involved in the development of laser induced CNV.International Journal of Ophthalmology 01/2013; 6(2):131-5. · 0.12 Impact Factor
Estradiol induces heparanase-1 expression and heparan
sulphate proteoglycan degradation in human endometrium
Xiulong Xu1,5, Jianchi Ding2, Geetha Rao1, Jikun Shen3, Richard A. Prinz1, Nasir Rana2and
1Departments of General Surgery, Rush University Medical Center, Chicago, IL, USA,2Institute for the Study and Treatment of
Endometriosis, Oak Brook, IL, USA,3Department of Surgery, University of Chicago, Chicago, IL, USA and4Rush Medical College,
Chicago, IL, USA
5To whom Correspondence should be addressed at: Department of General Surgery, Rush University Medical Center, 1653 W. Congress
Parkway, Chicago, IL 60612, USA. Tel: þ1 312 942 6623; Fax: þ1 312 942 2867; E-mail: email@example.com
BACKGROUND: This study seeks to determine whether estrogen is able to regulate the expression of heparanase-1
(HPR1) in human endometrium. METHODS: HPR1 expression and heparan sulphate (HS) deposition in the endome-
trium collected in various menstrual phases were analysed by immunohistochemical and immunofluorescence stain-
ing, respectively. HPR1 expression in the endometrial cells unexposed or exposed to estradiol was analysed by using
RT-PCR and luciferase reporter assay. HPR1 activity was analysed by using a novel enzyme-linked immunosorbent
assay (ELISA). Cell surface HS levels were analysed by flow cytometry. Serum HPR1 activity in women receiving
follicle-stimulating hormone (FSH) for IVF was measured by ELISA. RESULTS: HPR1 expression was rarely
detected in the endometrium in the early and mid-proliferative phases but was increased in the late proliferative
phase and in the secretory phases. HPR1 expression was negatively associated with HS in the basement membrane
(BM) of the endometrial glands. HPR1 gene expression, HPR1 promoter activity and HPR1 enzymatic activity
were increased in the endometrial cells when exposed to 17b-estradiol (E2), whereas cell surface HS levels showed
a decrease which could be blocked by PI-88, an HPR1 inhibitor. Serum HPR1 levels were increased in women
with moderately elevated blood estrogen levels after receiving FSH. CONCLUSIONS: HPR1 is differentially
expressed in the endometrium in different menstrual phases. Estrogen plays an important role in inducing HPR1
expression, subsequently leading to HS degradation on the endometrial cell surface and in the BM of the
Keywords: endometrium/estradiol/heparanase/heparan sulphate/ovarian hyperstimulation syndrome
Heparan sulphate proteoglyeans (HSPGs) are important
components of the cell surface, the extracellular matrix (ECM)
and the basement membrane (BM). At least 13 HSPG genes
from 5 distinct classes have been identified. They include
three pericellular HSPGs (perlecan, agrin and the hybrid
rimon andBernfield,2000).HSPGs comprise acoreprotein that
is covalently attached to a unique glycosaminoglycan chain
characterized by a linear array of alternating disaccharide
units (Stringer and Gallagher, 1997; Perrimon and Bernfield,
2000; Esko and Lindahl, 2001; Iozzo, 2001). Previous studies
have shown that the HSPG gene expression and the presence
of HS on HSPG protein cores can be regulated by steroid hor-
mones in the endometrium. For example, Russo et al. (2001)
reported that administration of 17b-estradiol (E2) leads to
increased expression of syndecan-3 in the uterus of ovari-
ectomized rats. Syndecan-3 is largely located in the epithelial
cells of glands and in the endometrial stroma as well as in the
smooth muscle cells of the myometrium. Potter and Morris
reported that syndecan-1 expression is decreased in the basolat-
the cycle progresses from metestrus toward estrus (Potter and
Morris, 1992). Paradoxically, Morris et al. (1988) showed that
E2can accelerate the turnover of HSPGs in the lysosomes of
murine endometrial epithelial cells. Though the significance of
HSPG expression and regulation in the endometrium remain
to be defined, it appears that the regulation of HSPG expression
and the status of HS in HSPGs by estrogen may have important
Heparanase-1 (HPR1) is an endoglycosidase that specifically
degrades HSPGs (Hulett et al., 1999; Vlodavsky et al., 1999;
Parish et al., 2001; Vlodavsky and Friedmann, 2001). HPR1
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Hum. Reprod. Advance Access published January 29, 2007
by guest on June 3, 2013
expression is up-regulated in a variety of malignancies and
plays an important role in tumour angiogenesis and metastasis.
HPR1 stimulates tumour angiogenesis by releasing the growth
factorssuch asfibroblast growthfactorandvascularendothelial
growth factor (VEGF) stored in the ECM (Elkin et al., 2001;
Goldshmidt et al., 2002; Vlodavsky et al., 2002; Elkin et al.,
2003; Edovitsky et al., 2004; Mikami et al., 2004; Cohen
of VEGF (Zetser et al., 2006) and cyclooxygenase-2 (Okawa
et al., 2005). HPR1 promotes tumour metastasis by degrading
HSPGs in the ECM and BM, allowing tumour cells to spread
to a distant site (Vlodavsky et al., 1999, 2002; Marchetti and
Nicolson, 2001; Vlodavsky and Friedmann, 2001; Edovitsky
as an adhesion molecule (Goldshmidt et al., 2003; Zetser et al.,
2003) and promote endothelial cell migration (Gingis-Velitski
et al., 2004a). Recent studies suggest that HPR1 is involved
in the pathogenesis of many other diseases such as diabetic
nephropathy (Maxhimer et al., 2005) and delayed type hyper-
sensitivity (Edovitsky et al., 2005).
HPR1 expression is also detected in several reproductive
cell types and may play an important role in tissue remodelling.
For example, HPR1 expression is detected in the extravillous
trophoblasts invading the decidua and in the endothelium
of fetal capillaries (Haimov-Kochman et al., 2002). HPR1
expression in these cells may facilitate embryo implantation
by promoting trophoblast cell invasion and tissue remodelling.
Indeed, Zcharia et al. (2004) demonstrated that transgenic mice
overexpressing HPR1 in all tissues under the control of a
chicken b-actin promoter have a significantly higher embryo
implantation rate than control mice. Further studies by these
investigators showed that pretreatment of mouse embryos
with recombinant HPR1 in vitro is able to increase the
implantation rate (Revel et al., 2005). It was not clear
whether HPR1 is also expressed in the endometrium and
whether HPR1 is involved in endometrial tissue remodelling
and in preparing the endometrium for embryo implantation
and subsequent angiogenesis. On the basis of a prior
study (Elkin et al., 2003) demonstrating the ability of
estrogen to induce HPR1 expression in breast cancer, we
hypothesize that HPR1 may be differentially expressed in
different menstrual phases in the endometrium, due to
regulation by steroid hormones. In the present study, we report
that HPR1 expression is increased in the endometrium in the
late-proliferative phase (LP) and plate secretory phases (LS),
and that estrogen is able to induce HPR1 expression and
HS degradation on the cell surface and in the BM of the
Materials and Methods
The use of human tissues was approved by the Rush University
Medical Center Institutional Review Board. Women with regular
menstrual cycles (28–30 days) of the reproductive ages were recruited
by the senior author (W.P.) from an IVF clinic as part of an infertility
evaluation. During laparoscopy, pelvic organs were examined for the
inflammatory disease, endometriosis and adhesions were not included
in the study. Samples of the uterine endometrium from 33 women
(mean age+ SD ¼ 32.7+ 7.2 years old, median age, 34 years old;
range 21–44 years old) were obtained with Novak’s curette from
the uterine fundus. No hormonal therapies were used in these 33
women during the cycle. The menstrual phases were classified as
early-proliferative (EP), mid-proliferative (MP), late-secretory (LP),
early-secretory (ES), mid-secretory (MS) and LS phases, on the
basis of the histological morphology and the date of menses as pre-
viously described (Noyes and Haman, 1953; Dmowski et al., 2001).
Part of each specimen was fixed immediately in 4% formaldehyde
and embedded in paraffin within 48 h in a single pathology laboratory.
Paraffin blocks of the uterine endometrial specimens were retrieved
for the study from the pathology laboratory repository.
To study the effect of estrogen in regulating HPR1 expression
in vivo, we tested whether increased blood estradiol levels correlated
with increased serum HPR1 levels. To address this, we analysed
HPR1 activity in the serum samples from seven women undergoing
controlled ovarian stimulation with follicle-stimulating hormone
FSH for IVF. The diagnoses for these infertile patients were
endometriosis (three), ovarian factor (two), male factor (one) and
tubal factor (one). The patients were treated with 150 to 600 units of
recombinant FSH (Follistim, Organon USA or GonalF, Serono,
USA) by s.c. injection until oocyte retrieval. The blood samples
were collected at various time points according to each individual’s
response. The dose of FSH was adjusted according to the ovarian
response to FSH stimulation, e.g. serum E2levels and the follicular
growth. The former was quantitfized by using an Immulite kit
(Diagnostic Products Corporation, LA, CA, USA); the later was
monitored by ultrasonography. After the hormonal assay, serum
samples were stored in a 280ºC freezer until assayed for HPR1
activity. Serum samples collected at multiple times during the IVF
cycle were used in this study.
Sections of endometrial specimens and an HPR1-positive pancreatic
adenocarcinoma that was included as a positive control (PC) were
de-waxed with xylene and rehydrated. Slides were heat-inactivated
in 10 mM sodium citrate (pH 6.0) in a microwave for 3 min. Cooled
slides were rinsed with PBS and then incubated with 1% H2O2in
methanol for 30 min at room temperature. Sections were blocked
with 5% normal goat serum in phosphate-buffered saline (PBS) for
30 min at room temperature, followed by 1 h incubation with an
anti-HPR1 rabbit antiserum (1:500 dilution) in PBS. This antiserum
was raised by immunizing a rabbit with a peptide containing the
amino acid residues from 273 to 290 of the 50 kDa HPR1 subunit
(Fairbanks et al., 1999). Normal rabbit serum was included as a nega-
tive control (NC). Slides were washed and then incubated with goat
anti-rabbit antibody–biotin conjugate (PharMingen, San Diego, CA,
USA) diluted at 1:300 in PBS with 5% human serum. Strepavidin–
horse-radish conjugate (Zymed, San Francisco, CA, USA) diluted at
1:200 in PBS with 5% normal human serum was added and incubated
for 45 min at room temperature. Colour development was done with
diaminobenzidine (DAB) substrate (Sigma, St. Louis, MO, USA) fol-
lowed by DAB enhancer (Vector Laboratories, Burlingame, CA,
USA). Slides were counterstained with Mayer’s haemotoxylin for
2-min, dehydrated and mounted. HPR1 expression was graded in a
blinded fashion by two investigators (J.S. and X.X.) in this study.
Negative HPR1 expression was defined as no HPR1 or weak
signal detected in ,10% of the stromal or glandular cells. Positive
HPR1 expression was defined as HPR1 signal in at least 10% of
cells with moderate or strong intensity in either the stromal or gland-
X.Xu et al.
Page 2 of 11
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Immunofluorescence analysis of HS deposition
dehyde. The slides were incubated with an anti-HS mAb (Clone
HepSS, Seikagaku Corp., Chuo-ku, Tokyo, Japan) at a concentration
of 5 mg ml21at room temperature for 30 min. The same concentration
of normal mouse IgM was used as a negative control. The slides were
washed three times with PBS, followed by incubation with a goat anti-
mouse IgM conjugated with FITC (1:50) (ICN Biomedicals, Aurora,
OH, USA) and then washed and incubated with rabbit anti-goat IgG
conjugated with FITC (1:50) (ICN Biomedical, Aurora, OH). The
slides were washed and sealed with 50% glycerin in PBS containing
anti-fade reagent 1,4-diazabicyclo(2,2,2)octane
(25 mg ml21). HS expression was examined under fluorescent
microscopy. Negative HS staining was defined as the absence of fluor-
escence signal in the BM in .90% of the glands. Positive HS staining
was defined as the presence of fluorescence signal in the BM in .10%
of the glands.Thepictureswere taken witha digitalcamera attachedto
a Nikon Eclipse TE200 fluorescence microscope.
Endometrial cell culture and stimulation with E2
Approximately 200 mg of fresh tissue in the LP or ES phase were
minced into small pieces (1–2 mm3) and washed in fresh medium to
remove mucus or debris. Tissue fragments were then digested in the
(200 units ml21) at 378C for 45 min by stirring. At the end of incu-
bation, cell clumps were further mechanically dispersed by aspiration
with a Pasteur pipette. The mixture of single stromal cells and the
large clumps of epithelium were washed twice with Hank’s balanced
salt solution (Pharmacia, Piscataway, NJ, USA). The stromal and epi-
thelialcells were notfurtherseparated,sinceourimmunohistochemical
(IHC) analysis revealedthat HPR1 expression was detected in both cell
types. Primary endometrial cells were grown in RPMI 1640 sup-
plemented with 5% charcoal/dextran-treated fetal bovine serum
(Hyclone, Logan, Utah). After incubation for 24 h, the unattached
float cells were removed. In some experiments, the medium was
coal/dextran-treated fetal bovine serum. There was no significant
difference in the experimental results conducted with the medium con-
taining phenol red or no phenol red. The monolayers of endometrial
cells from a 72-h primary culture were trypsinized and seeded in
6-well plates. Cells were left unstimulated or stimulated with various
concentrations of E2. After incubation for 48 h, the conditioned media
were collected and spun down at 48C, 15 000 g for 15 min. The super-
natants were collected and stored at 2808C until use. The cells were
washed three times with PBS and then detached by incubation with
5 mM EDTA in PBS for 5 min. The cells were lysed in HPR1 assay
buffer [0.1 M sodium acetate, pH 5.0, 0.1 mg ml21bovine serum
albumin (BSA), 0.01% Triton X-100, 0.5 mM phenylmethylsulphonyl
fluoride (PMSF), and 10 mg ml21leupeptin and aprotinin each] fol-
lowed by three quick freeze and thaw cycles. The cell lysates were
spundownat48C,15 000 gfor15 min.Thesupernatantswerecollected
and analysed for protein concentration by using a Protein Assay kit
(Bio-Rad, Hercules, CA, USA). Cell lysates were analysed for HPR1
activity by using an enzyme-linked immunosorbent assay (ELISA)
method as described below.
(100 mg ml21) and DNase
centrationsofE2for24 h.Cellswerelysedin1 mlTRIzolandRNAwas
extracted by following the manufacturer’s instruction (Life Technol-
ogies). RNA concentration was quantified by ultraviolet absorption.
After reverse transcription of 500 ng total RNA with oligo(dT)
priming, the resulting single stranded cDNA was amplified using Taq
DNA polymerase (Life Technologies). Oligonucleotides HPR-3
(50-TTCGATCCCAAGAAGG-AATCAAC-30) and HPR-4 (50GTAG
TGATGCCATGTAACTGAAT-C-30) were used for amplifying a
587 bp heparanase cDNA fragment. Oligonucleotides 50TGAAGGTC
AG-TCCTTCCACG-30were used to amplify a 527 bp GAPDH
cDNA fragment. The PCR conditions were an initial denaturation of
4 min at 948C and subsequent denaturation for 45 s at 948C, annealing
for 1 min at 558C and extension for 1 min at 728C (32 cycles). Aliquots
of10 mlPCRproductswereseparated by 1.5%agarose gel electrophor-
esis and visualized by ethidium bromide staining.
Luciferase reporter gene expression
Primary endometrial cells were prepared and grown in RPMI 1640 sup-
plemented with 5% charcoal/dextran-treated fetal bovine serum in a
T-75 flask. Cells were harvested and washed twice with PBS. Cells
(5 ? 106cells per sample) were transfected with 8 mg of plasmid con-
moter fragment (pGL3/HPR-0.3 and pGL3/HPR-3.5) and 2 mg of
pCMV/SPORT containing the b-galactosidase gene as an internal
control by electroporation with 300 V and 900 mCi in a Gene Pulser II
(Bio-Rad, Hercules, CA, USA). Luciferase reporter constructs have
been previously described (Jiang et al., 2002). A plasmid containing
the luciferase reporter gene without an HPR1 promoter (pGL3/Basic)
was included as an NC. Cells were then washed and seeded in a
of E2. After incubation for 48 h, cells were harvested, and cell lysates
were prepared. In some experiments, cells were also transfected with
FuGENE6 transfection reagent (Roche Applied Science, Indianapolis,
IN, USA), following the manufacturer’s instruction. Similar results
were obtained. The luciferase activity was quantified by using the luci-
ferin substrateand read ina TECAN plate reader (Phenix Research Pro-
ducts, Hayward, CA, USA). The relative light unit in each sample was
normalized against the b-galactosidase activity measured by a colori-
metric assay, as previously reported (Eustice et al., 1991). The means
Flow cytometric analysis
Endometrial cells from the primary cell culture in T-75 flasks were
seeded in 6-well plates and stimulated with E2 (1 ? 1028or
2 ? 1027M) for 24 h in the absence or presence of PI-88
(50 mg ml21). Single cell suspensions of endometrial cells were pre-
pared by using Cell Dissociation Solution. Cells (5 ? 105per
sample) were stained by incubation for 30 min at 48C with an
anti-HS mAb (0.5 mg per sample) (clone HepSS, Seikagaku Corp.,
Chuo-ku, Tokyo, Japan) or mouse IgM as an isotype control. Cells
were incubated with FITC-labelled goat anti-mouse IgM (5 ml per
sample) for 30 min at 48C, followed by rabbit FITC-labelled anti-goat
IgG (5 ml per sample) for 30 min at 48C. Cell surface HS expression
was analysed in a Becton Dickson flow cytometer.
HPR1 activity assay
HPR1 activity in the serum samples, cell lysates and supernatants of
viously described (Quiros et al., 2006). Briefly, Matrigel (BD Bio-
sciences, San Diego, CA, USA), an artificial BM which contains
abundant HSPGS, was dissolved in ice-cold PBS (0.1 M, pH 7.4)/car-
bonate-buffered saline (0.1 M, pH 9.6) (volume:volume, 50:50) at a
concentration of 20 mg ml21and used to coat ELISA plates (25 ml
per well) at 48C overnight. The plates were then washed three times
with PBS containing 0.05% Tween-20 and blocked with 5% BSA in
E2induces HPR1 expression in endometrium
Page 3 of 11
by guest on June 3, 2013
PBS at room temperature for 1 h. Serum samples were diluted at 1:5 in
HPR1 assay buffer (0.1 M sodium acetate, pH 5.0, 0.1 mg ml21BSA,
0.01% Triton X-100, 0.5 mM PMSF, and 10 mg ml21leupeptin and
aprotinin each). The supernatants of cultured endometrial cells were
premixed with 10? HPR1 assay buffer. Diluted serum samples, super-
natant and cell lysates (25 ml per well) were added to each well and
incubated at 378C overnight. After a wash, anti-HS-specific mAb
(clone HepSS, Seikagaku Corp., Chuo-ku, Tokyo, Japan) (1:1000,
diluted in PBS containing 5% BSA) was added and incubated at
room temperature for 1 h. After washing, horse-radish peroxidase-
conjugated goat anti-mouse IgM antibody (1:2000 diluted in PBS
with 5% BSA) was added and incubated at room temperature for 1 h,
followed by addition of 50 ml 2,2-azino-bis-(3-ethylbenzthiazoline-
6-sulfonic acid substrate. The OD405 absorbance was read in an
ELISA plate reader (Bio-Rad, Hercules, CA, USA). HPR1 activity in
serum samples was calculated on the basis of a standard curve of
serially diluted purified platelet HPR1 (starting at 1:200) at a concen-
tration of 1 ml HPR1 with the activity of degrading 0.133 mg HS per
hour at 378C in HPR1 buffer. HPR1 purification and characterization
from human platelets were conducted, as previously reported (Ihrcke
et al., 1998). HPR1 activity was designated as per 100 units capable
of degrading 1 ng HS at 378C h21in HPR1 buffer.
x2test was used to analyse the significance of a difference in the fre-
quency of HPR1 expression in the proliferative and secretory phases.
Fisher’s exact test was used to analyse the relationship of HPR1
expression with the deposition of HS in the BM. Spearman rank
order correlation test was used to analyse the correlation between
serum HPR1 activity and blood E2levels. A P-value , 0.05 was
considered as statistically significant. All statistics were conducted
by using the SigmaStat3 software (Richmond, CA, USA).
IHC analysis of HPR1 expression in the endometrium
We first conducted IHC analysis to determine whether HPR1
expression differed in the endometrial tissue at the different
phases of the menstrual cycle. As shown in Figure 1, HPR1
was neither expressed in glandular epithelial cells nor in the
stromal cells in two endometrial specimens collected in
the EP and MP phases. However, HPR1 was detected in the
stromal and glandular cells in the endometrial specimens
collected in the LP and ES, MS and LS phases. Approximately
80% of the specimens expressed HPR1 in both the stromal cells
and glandular epithelia in the functionalis layers with com-
parable intensity. A normal rabbit serum was used as a NC;
non-specific signals were not observed in this endometrial
specimen collected in the LP phase. A HPR1-positive pancrea-
tic adenocarcinoma was included as a PC; strong signals were
observed in the tumour cells.
Among 33 endometrial specimens analysed, we found that
HPR1 expression was detected in one of seven endometrial
specimens (14%) in the EP and MP phases each. HPR1 was
detected in three of six specimens taken in the LP phases and
in 10 of 13 specimens during the secretory phases (Table I).
Fisher’s exact test revealed that HPR1 expression in the
secretory phases (HPR1 positive in 10 of 13 specimens) was
significantly higher than in the LP phases (HPR1 positive in
Figure 1. Immunohistochemical staining (IHC) of heparanase-1 (HPR1) expression. HPR1 expression was not present in endometrium in the
early- and mid-proliferative (EP and MP) phase, but was strongly present in both the stromal and glandular cells in the endometrium in the
late proliferative (LP), and early- mid- and late-secretory (ES, MS and LS) phases. Normal rabbit serum was included as a negative control
(NC). A HPR1-positive pancreatic adenocarcinoma was included as a positive control (PC) for IHC staining.
X.Xu et al.
Page 4 of 11
by guest on June 3, 2013
2 of 14 specimens) (P ¼ 0.002), but was not significantly
higher than in the LP phase (HPR1 positive in three of six
specimens) (P ¼ 0.32).
Detection of heparan sulphate proteoglycan in the BM
We next tested whether increased HPR1 expression in the
endometrium led to the degradation of HS in the BM of the
HPR1-positive and HPR1-negative endometrial tissues was
analysed by using immunofluorescence (IF) staining with a
monoclonal antibody specific for HS. As shown in Figure 2,
HS deposition was present in the BM of an HPR1-negative
endometrial specimen collected in the MP phase (A) but was
absent in the BM of a HPR1-positive endometrium collected
in the MS phase (B). In some specimens, HS signal was also
present in the nuclear membrane. Normal mouse IgM was
included as an NC and no signal was observed (C). We analy-
sed the relationship between HPR1 expression and the deposi-
tion of HS. As shown in Table II, 7 of 10 HPR1-positive
endometrial specimens did not exhibit HS deposition in the
BM, whereas 10 of 12 HPR1-negative endometrial specimens
had intact HS deposition. Statistical analysis revealed that
HPR1 expression was negatively associated with the presence
of HSGP in the endometrium (P ¼ 0.036), suggesting that
HPR1 expression is responsible for the degradation of HS in
the BM of the endometrium.
expression in the BM of
Induction of HPR1 expression in endometrial cells
Recently, Elkin et al. (2003) reported that E2is able to induce
HPR1 expression in MCF7 cells, a breast cancer cell line. We
hypothesized that increased HPR1 expression in the LP and LS
phases may be due to the rising estrogen levels. We first
conducted a semi-quantative RT–PCR to test whether E2
was able to induce HPR1 mRNA expression in endometrial
cells. As shown in Figure 3A, E2 at the concentration of
1029M dramatically induced HPR1 mRNA expression,
whereas continued increase in E2concentration was relatively
less effective in inducing HPR1 expression. To further test
whether E2-induced HPR1 gene expression was due to
increased promoter activation, we conducted luciferase
reporter gene assays in primary endometrial cells using
four luciferase reporter constructs. As shown in Figure 3B,
E2had no effect on the luciferase activity in the cells tran-
fected with the empty vector or the vector containing a
0.3-kb HPR1 promoter fragment with the luciferase gene
(pGL3/HPR1-0.3). However, in comparison with untreated
cells, E2 at the concentration of 1 ? 1029M consistently
increased luciferase activity by ?60% in the cells trans-
fected with plasmids containing the luciferase reporter
gene driven by a 3.5-kb HPR1 promoter fragment (pGL3/
HPR1-3.5). We next tested whether induction of HPR1 pro-
moter activity by E2corresponded to the induction of HPR1
mRNA expression. As shown in Figure 3C, E2effectively
induced expression of a 3.5-kb HPR1 promoter-driven luci-
ferase gene, however at the concentration of 1 ? 1029M, E2
was slightly more effective in inducing HPR1 promoter
activity than when it was used at 1 ? 1028and 1 ? 1027M.
To further confirm the ability of E2 to induce HPR1
expression, we conducted an enzymatic assay to test whether
HPR1 activity in the cell lysates and in the supernatant of the
endometrial cells exposed to E2was increased. As shown in
Figure 3D, HPR1 activity was increased in the cell lysates of
the endometrial cells treated with E2at 10210, 1029, 1028or
1 ? 1027Mby35,48,63,and31%,respectively.HPR1activity
with E2at 10210, 1029, 1028or 1 ? 1027M by 34, 43, 77, and
41%, respectively. E2used at the concentration 1 ? 1028M
cate that E2used at very high concentrations (.1 ? 1028M)
was less effective in inducing HPR1 expression.
Induction of cell surface heparan sulphate degradation by E2
We next tested whether induction of HPR1 expression by E2
in endometrial cells led to the degradation of cell surface HS.
Endometrial cells were left unstimulated or stimulated with
Table I. Heparanase-1 (HPR1) expression in the endometrium during the
EP, early-proliferative; MP, mid-proliferative; LP, late-proliferative; ES,
early-secretory; MS, mid-secretory; LS, late-secretory. HPR1 expression in
the proliferative phase versus the secretory phase, P ¼ 0.005.
Figure 2. Immunodetection of heparan sulphate proteoglycan (HSPG) in the endometrial tissue (A) HS deposition was present in the basement
membrane (BM) of an endometrial gland from an HPR1-negative endometrium specimen collected in the EP phase. (B) No HS signal was present
in the BM of the endometrial glands in an HPR1-positive specimen collected in the MS phase. (C) Mouse IgM was included as isotype control,
showing no non-specific staining.
E2induces HPR1 expression in endometrium
Page 5 of 11
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E2(1 ? 1028or 2 ? 1027M) in the absence or presence of
PI-88 (50 mg ml21) for 48 h. As shown in Figure 4A, cell
surface HS was detected at modest levels in untreated cells.
However, cell surface HS levels were slightly decreased in
the endometrialcells exposed
(Figure 4B). Reduction of cell surface HS levels was much
more pronounced when the endometrial cells were treated
with E2 at 1 ? 1028M (Figure 4C). To confirm that
E2-induced cell surface HS degradation was mediated by
increased HPR1 expression, we tested whether PI-88, a
novel HPR1 inhibitor, was able to restore cell surface HS
levels in E2-treated cells. As shown in Figure 4E, PI-88
(50 mg ml21) dramatically increased cell surface HS levels
in E2-treated endometrial cells compared with those in
E2-treated cells without PI-88 (Figure 4C). PI-88 also slightly
increased cell surface HS levels in untreated cells (compare
Figure 4D and A). These observations strongly suggest that
the decrease in cell surface HS levels in E2-treated glandular
cells is mediated by accelerated degradation due to increased
at 2 ? 1027M
Serum heparanase-1 levels in in vitro fertilization patients
We next tested whether moderately increased E2levels led to
an optimal increase of serum HPR1 activity, whereas excessive
E2levels in blood led to a lesser increase of serum HPR1
activity. To address this, we analysed serum HPR1 levels in
seven women treated with FSH for IVF. As shown in
Figure 5A–G, continued FSH treatment led to a linear increase
of blood E2levels in all five of seven patients. Interestingly,
serum HPR1 levels were maximally increased in all seven
patients when their blood E2levels ranged between 300 and
900 pg ml21(1.1–3.3 ? 1029M). It should be noted that E2
concentration in the uterus is ?10-fold higher, e.g. ?at
1028M (Bulun et al., 2005). Further increases in blood E2
levels (.900 pg ml21) did not lead to further increase in
serum HPR1 activity, but rather a decrease in serum HPR1
activity in all patients, compared with that in the samples
with optimal blood E2levels. To further examine the role of
estrogen in regulating HPR1 expression, we analysed the
relationship between serum HPR1 activity and estradiol
levels in 18 samples whose serum estradial levels were
,900 pg ml21. As shown in Figure 5H, serum HPR1 activity
was very low in 10 samples whose serum estradiol levels
were ,215 pg ml21but was dramatically increased in eight
samples whose serum estradiol levels were ,215 pg ml21.
The spearman rank order correlation test revealed that serum
HPR1 activity in 18 samples from 7 patients correlated well
with blood E2levels (P , 0.001).
Our present study provides several lines of evidence that
estrogen is able to regulate HPR1 expression in human endo-
metrium in vitro and in vivo: (i) E2was able to activate the
HPR1 promoter and to induce HPR1 gene expression in the
endometrial cells; (ii) Up-regulation of HPR1 expression by
E2led to accelerated degradation of cell surface HS, which
was blocked by a HPR1 inhibitor; (iii) Serum HPR1 levels
were increased when blood estrogen levels were moderately
elevated after FSH administration; (iv) HPR1 expression
was increased in the endometrial specimens taken in the LP
phase when blood estrogen levels are increased. It should be
noted that because of the small number of samples, we
were unable to find that HPR1 expression was significantly
higher in the late than in the EP and MP phases in normal
endometrium, but it was indeed significantly increased in
the endometrium from women with endometriosis (X. Xu,
unpublished data). The ability of E2 to induce HPR1
expression in the endometrial cells is in agreement with a
prior observation made by Elkin et al. (2003) in MCF-7
cells, a breast cancer cell line. Intriguingly, the optimal con-
centrations of E2 to induce HPR1 expression in both the
breast cancer cell line and the endometrial cells were approxi-
mately between 1028and 1029M. Higher E2concentrations
(.1028M) were less effective in inducing HPR1 expression
and subsequent cell surface HS degradation. Consistent with
these in-vitro observations, our clinical study showed that
serum HPR1 levels were increased in the IVF patients with
moderately increased blood E2levels (up to 900 pg ml21or
3.3 ? 1029M); however, a further increase of blood E2
levels did not lead to a further increase of serum HPR1
Although our present study and the study by Elkin et al.
(2003) have provided strong evidence that HPR1 expression
can be regulated by estrogen, the underlying molecular mech-
anisms are not fully understood. RT–PCR and luciferase
reporter gene assays suggest that E2-induced HPR1 expression
is at the transcriptional level. Numerous putative estrogen-
responsive elements (EREs) are found to be present in the
HPR1 promoter (Elkin et al., 2003). However, it is not clear
whether HPR1 gene transactivation is mediated by direct
binding of estrogen receptors (ERs) to these ERE sites. Pre-
vious studies have shown that estrogen can regulate the
expression of low-density lipoprotein receptor and several
others genes with a GC-rich promoter, including VEGF, reti-
noic receptor a, TGF-a and progesterone receptor (PR),
through the interaction of ER and the transcription factor Sp1
(see review by Bjornstrom and Sjoberg, 2005). Moreover,
estrogen can activate Raf-1 kinase in an ER-independent
manner (Singh et al., 2000). Raf kinase activation induces
the expression of Egr-1 transcription factor (Pratt et al.,
1998) and activates the Ets family transcription factors such
as Ets-1 (Lincoln et al., 2003). Interestingly, all of these
Table II. HPR1 expression and heparan sulphate (HS) deposition in the
basement membrane (BM)
NumberHS Positive (%)
HS positivity in HPR1-positive specimens versus HS positivity in HPR1-
negative endometrium, P ¼0.036.
X.Xu et al.
Page 6 of 11
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three families of transcription factors, Sp1, Egr-1 and Ets, have
been shown to play an important role in regulating HPR1 gene
expression (Jiang et al., 2002; de Mestre et al., 2003, 2005; Lu
et al., 2003; Ogishima et al., 2005). Therefore, it is likely that
these transcription factors may act in concert with ER to
mediate estrogen–induced HPR1 expression.
Recent in-depth studies revealed that HPR1 has multiple
of normal and tumour tissues but also in tumour cell invasion.
In addition, HPR1 can function as an adhesion molecule (Gold-
shmidt et al., 2003; Zetser et al., 2003) and contributes to cell
migration (Gingis-Velitski et al., 2004a). Numerous studies
of both the human and bovine placenta (Dempsey et al., 2000;
Kizaki et al., 2001, 2003; Haimov-Kochman et al., 2002) and
that HPR1 expression may assist trophoblastic cell invasion
into the endometrium and promote angiogenesis. In support of
this, Zcharia et al. (2004) reported that the embryo implantation
Figure 3. Induction of HPR1 expression by E2. (A) Induction of HPR1 mRNA by E2. Second passage endometrial cells were allowed to reach
80% confluence and then stimulated with the indicated concentration of E2. After 24 h, the cells were analysed for HPR1 expression by RT–PCR.
(B) Activation of HPR1 promoter activity by E2. Second passage endometrial cells were transfected with HPR1 promoter-driven luciferase repor-
ter plasmids and after 48 h, analysed for luciferase activity. The results from two independent experiments are shown as the mean+SD of
triplicate test in each experiment. (C) Dose–response of HPR1 promoter-driven luciferase reporter gene expression. Second passage endometrial
cells were transfected with a 3.5-kb HPR1 promoter-driven luciferase reporter construct and then left unstimulated or stimulated with indicated
concentrations of E2. After 48 h, cells were analysed for luciferase activity. (D and E) Increased HPR1 activity in the cell lysates and supernatants
of endometrial cells exposed to E2. Endometrial cellswere treated with the indicated concentrations of E2for 48 h. HPR1 activity in the cell lysates
(D) and supernatants (E) were then measured by enzyme-linked immunosorbent assay (ELISA).
E2induces HPR1 expression in endometrium
Page 7 of 11
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implantation rate in HPR1-transgenic mice may also be in part
treatment of mouse embryos with recombinant HPR1 in vitro is
able to increase the embryo implantation rate (Revel et al.,
2005). Our present study demonstrated that HPR1 expression
was increased in the endometrium in the LP phase and remained
at high levels throughout the luteal phases. Though the physio-
dation in the endometrium remains unclear, we speculate that
HPR1may play an importantroleinendometrial tissueremodel-
and facilitating trophoblast cell invasion as well as angiogenesis.
The observations that HPR1 expression can be maximally
induced by an optimal concentration of estrogen may have
potential physiological significance. Zcharia et al. (2004)
reported that overexpression of HPR1 in transgenic mice
leads to increased embryo implantation. However, the survival
rate of implanted embryos is significantly lower in HPR1-
transgenic mice than in the wild-type mice (Zcharia et al.,
2004), probably due to HPR1-mediated excessive angiogenesis
in fetal tissues and/or in the placenta. During pregnancy, blood
E2levels range between 1000 and 5000 pg ml21in the first
trimester, 5000 and 15 000 pg ml21in the second trimester
and 10 000 and 40 000 pg ml21in the third trimester. Thus,
the suboptimal induction of HPR1 expression by estrogen at
very high concentrations in the second and third trimesters
may allow the fine-tuning of angiogenesis and avoid the detri-
mental effect of excessive angiogenesis in the placenta and/or
in the fetal tissue.
During the preparation of this article, Kodama et al. (2006)
reported that HPR1 expression was not detected in 11 normal
endometrial specimens in the EP phase but was detected in 4
specimens in the LP phase and that HPR1 was expressed at a
moderate level in the secretory phase. Similar to these findings,
we found that HPR1 was rarely expressed in normal endo-
metrial specimens in the EP and MP phases but was detected
in three of six endometrial specimens in the LP phase.
Because of the small number of samples, we were unable to
find a significant difference in HPR1 expression in the endome-
trium between the EP and LP phases, and between MP and LP
phases. However, we found that HPR1 expression was detected
in the majority of endometrial specimens in the secretory
phases. In particular, we found that HPR1 expression was
detected in all four endometrial specimens in the LS phase,
while blood E2levels should have already declined in this
period. It is not clear whether there is alapse for HPR1 turnover
in the endometrial tissue. Alternatively, HPR1 expression may
be regulated by other hormones. In support of this notion, our
unpublished data indicate that progesterone was able to weakly
induce HPR1 expression and HS degradation in endometrial
cells. Progesterone may regulate HPR1 expression with a
mode of action similar to estrogen, e.g. via interaction of PR
with Sp1 (Owen et al., 1998) and other GC-rich binding tran-
scription factors such as basic transcription element binding
protein, which are highly expressed in the endometrium
(Zhang et al., 2001, 2002, 2003). Alternatively, pogesterone
can also activate the Src tyrosine kinase pathway (Edwards,
2005), leading to the activation of Raf kinase and Ets trans-
cription factor, and the induction of Egr-1. Nevertheless, our
Figure 4. E2induces endometrial cell surface HS degradation. Endometrial cells were incubated in the absence or presence of indicated concen-
trations of E2and/or PI-88 for 48 h. Cells were then analysed for cell surface HS expression by staining with an HS-specific monoclonal antibody
followed by fluorescence-activated cell sorter. The black line is a mouse IgM control; the green line is an anti-HS IgM. (A) Untreated control;
(B) cells treated with E2(2 ? 1027M); (C) cells treated with E2(1 ? 1028M); (D) cells treated withPI-88 (50 mg ml21); (E) cells treated with E2
(1 ? 1028) plus PI-88 (50 mg ml21). The experiments were repeated twice with similar results.
X.Xu et al.
Page 8 of 11
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in-vitro and clinical studies, along with that of Kodama et al.
(2006), strongly suggest that HPR1 expression in the endo-
metrial cells can be up-regulated by E2.
The ability of E2to induce HPR1 and HS degradation in the
endometrial cells is in line with a prior study (Morris et al.,
1988) showing that estrogen is able to increase the turnover
of HSPGs in the lysosomes of the murine uterine epithelial
cells in which HPR1 is localized and processed to become an
enzymatically active latent enzyme (Gingis-Velitski et al.,
2004b; Zetser et al., 2004). Consistent with this, analysis of
Figure 5. Comparison of serum HPR1 and estrogen levels in IVF patients receiving follicle-stimulating hormone (FSH). (A–G) Blood samples
were collected from seven women (each graph represents the data from an individual patient) at the indicated days after starting FSH treatment.
Serum HPR1 activity was analysed by using ELISA (left y-axis) and presented as a bar graph. Serum estradiol levels (right y-axis) were presented
as a linear graph. Serum estradiol levels in the range of 300–900 pg ml21(shadowed area) produced the highest serum HPR1 activity in all seven
patients. (H) Correlation of serum HPR1 levels with serum E2levels. Estrogen levels in 18 serum samples from 7 patients with serum E2levels
,900 pg ml21were plotted against serum HPR1 activity. Spearman rank order test revealed that serum HPR1 activity correlated with serum E2
levels (P , 0.001).
E2induces HPR1 expression in endometrium
Page 9 of 11
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HS revealed that HPR1 expression was associated with the lack
of HS deposition in the BM of the endometrial glands, further
suggesting that HPR1 plays a critical role in degrading HS.
However, we found that HS deposition was detected in
the BM in a few HPR1-positive endometrial specimens. This
could be due to incomplete degradation of HS in the BM of
the endometrial glands or due to an E2-mediated increase of
HSPG biosynthesis (Russo et al., 2001), which may offset
the degradation of HS side chains mediated by HPR1. Never-
theless, the observations of increased turnover of HSPGs in
the endometrial epithelial cells may reflect changes associated
with blastocyst attachment and invasion of the endometrium.
The ovarian hyperstimulation syndrome (OHSS) is an iatro-
genic complication manifested by massive ovarian enlarge-
ment, extravascular fluid accumulation, intravascular volume
depletion, renal failure and hypovolemic shock (Kaiser,
2003; Delbaere et al., 2004; Budev et al., 2005). This compli-
cation often occurs following the administration of FSH in
women undergoing IVF. The molecular pathogenesis of this
syndrome is poorly understood. Numerous studies suggest
that elevated plasma VEGFlevels are responsible for triggering
the onset of OHSS (Geva and Jaffe, 2000). Interestingly, a
recent study by Zester et al. (2006) demonstrated that HPR1
is able to induce the expression of VEGF. In addition, HPR1
is able to directly damage the endothelial barrier by degrading
pericellular HSPGs of the endothelial cells. Edovitsky et al.
(2005) reported that vessel permeability and extravasation of
leukocytes and plasma proteins are increased at the site of
delayed-type hypersensitivity-associated inflammation as a
result of increased HPR1 expression in the endothelium. Con-
sistently, Negrini et al. (2005) reported that an intravenous
injection of a bolus of heparanase into rabbits leads to HS
degradation and development of oedema in the lungs. Our
present study shows that serum HPR1 levels in IVF patients
were elevated particularly when blood estrogen levels were
moderately increased. On the basis of these findings, we
propose that the persistence of high serum HPR1 levels in
women with moderately elevated estrogen levels may be a
key to triggering the onset of OHSS. Elevated serum HPR1
levels may increase the vessel permeability by directly
disrupting the BM of the endothelium and/or indirectly by
stimulating the production of VEGF. Further studies are
under the way to determine whether IVF patients who
develop OHSS have persistently high serum HPR1 and/or
In summary, this study provides compelling evidence that
HPR1 expression in the endometrium is regulated by E2and
that HPR1 expression leads to HS degradation on the cell
surface and in the BM of the endometrial glands.
This work was supported in part by Department of General Surgery at
Rush University Medical Center and by the Institute for the Study and
Treatment of Endometriosis, Oak Brook, IL, USA. We are grateful to
Dr Robert L. Heinrikson (Pharmacia & Upjohn, Inc., Kalamazoo, MI,
USA) for kindly providing rabbit anti-HPR1 serum and to Progen
Industrials Limited (Queensland, Australia) for kindly providing
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Submitted on August 7, 2006; resubmitted on September 22, 2006; resubmitted
on November 11, 2006; accepted on November 29, 2006
E2induces HPR1 expression in endometrium
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